Chapter 1 Introduction 1 1. INTRODUCTION 1.1 Lignocellulose –A valuable resource Lignocellulose a major structural component of woody plants and non-woody plants such as grass and represent a major renewable source which is available in large amount (~5500 tons) per year. The chemical properties of the components of lignocelluloses like lignin, cellulose and hemicelluloses make them a substrate for enormous biotechnological value (Malherbe and Cloete, 2003). Large amount of lignocellulosic waste is generated through forestry and agricultural practices, paper- pulp industries, timber industries and many agro-industries pose an environmental pollution problem. Much of the lignocellulosic waste is often disposed by biomass burning, which is not restricted to the developing countries alone, but is considered as a global phenomenon (Levin, 1996). However, the huge amount of plant biomass considered as a “waste” can potentially be converted in to various different valuable products, including biofuels, chemicals, and cheap energy source for fermentation and improved animal feeds. Moreover, different biocomposites (biodegradable composites) have been produced by incorporation of lignocellulose fillers in to biodegradable aromatic polyester, polybutylene adipate-co teraphthalate, which are a by-product of an industrial fractionation process based on wheat straw and other agro industrial wastes (Avèrous and Le Digabel, 2006). The bioconversions of lignocellulosic material to useful high value products normally require multistep processes and can be achieved by mechanical, chemical or biological treatment. Exploration of an efficient and green oxidation technologies using ligninolytic enzyme can serve as an environmentally benign alternative, for the treatment of lignocellulosic material which include (i) Enzymatic pretreatment (Grethlein 1984 and Grethlein and Converse, 1991). (ii) Hydrolysis of the polymer to produce readily metabolizable molecules (e.g. hexose or pentose sugars). (iii) Bioutilization of these molecules to support microbial growth or to produce economically viable bioproducts. Figure 1.1 summarizes the generalized process stages in lignocelluloses bioconversion in to value added products.
Transcript
Chapter 1 Introduction
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1. INTRODUCTION
1.1 Lignocellulose –A valuable resource Lignocellulose a major structural component of woody plants and non-woody plants
such as grass and represent a major renewable source which is available in large
amount (~5500 tons) per year. The chemical properties of the components of
lignocelluloses like lignin, cellulose and hemicelluloses make them a substrate for
enormous biotechnological value (Malherbe and Cloete, 2003). Large amount of
lignocellulosic waste is generated through forestry and agricultural practices, paper-
pulp industries, timber industries and many agro-industries pose an environmental
pollution problem. Much of the lignocellulosic waste is often disposed by biomass
burning, which is not restricted to the developing countries alone, but is considered as
a global phenomenon (Levin, 1996). However, the huge amount of plant biomass
considered as a “waste” can potentially be converted in to various different valuable
products, including biofuels, chemicals, and cheap energy source for fermentation
and improved animal feeds. Moreover, different biocomposites (biodegradable
composites) have been produced by incorporation of lignocellulose fillers in to
biodegradable aromatic polyester, polybutylene adipate-co teraphthalate, which are a
by-product of an industrial fractionation process based on wheat straw and other agro
industrial wastes (Avèrous and Le Digabel, 2006).
The bioconversions of lignocellulosic material to useful high value products normally
require multistep processes and can be achieved by mechanical, chemical or
biological treatment. Exploration of an efficient and green oxidation technologies
using ligninolytic enzyme can serve as an environmentally benign alternative, for the
treatment of lignocellulosic material which include (i) Enzymatic pretreatment
(Grethlein 1984 and Grethlein and Converse, 1991). (ii) Hydrolysis of the polymer to
produce readily metabolizable molecules (e.g. hexose or pentose sugars). (iii)
Bioutilization of these molecules to support microbial growth or to produce
economically viable bioproducts. Figure 1.1 summarizes the generalized process
stages in lignocelluloses bioconversion in to value added products.
Chapter 1 Introduction
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Figure 1.1: Generalized process stages in lignocelluloses bioconversion in to value added bioproducts (Howard et al, 2003)
All lignocelluloses material has structural similarities and to understand its biological
potential it is very crucial to know the complex structure of the cell walls. Due to their
highly lignified cell walls. The structures of cell wall are resistant to attack by
microorganisms and associated enzyme systems.
1.2 Structural features of lignocellulosic material Wood, grasses, and most of the plant litter represent a major part of the biomass in
nature and are collectively called lignocelluloses (Kuhad et al, 1997).Lignocellulose
is mainly composed of cellulose, hemicelluloses and lignin (Sjöström, 1993).
Amount of cellulose and lignin present are 2.5x1011tons and 2-3x1011tons respectively
on the earth representing 40% and 30%, while other polysaccharide comprising 26%
of organic matter carbon (Fengel and Wegener 1989, Argyropoulos and Menachem
1997). They are not uniformly distributed in the plant cell wall (Figure 1.2); the S2-
layer of the secondary wall has the highest percentage of cellulose, and the middle
lamella has the highest percentage of lignin, but all three compounds can be found in
every cell wall layer (Sjöström, 1993 and Kuhad et al, 1997). Gramineous plants have
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more variation than woody plants. In addition, some grasses contain considerable
amount of pectin in the middle lamella, where as wood contains only small quantities
of extractives, inorganic compounds and pectin compounds (Fengel and Weneger
1989; McDougall et al, 1993; Kuhad et al, 1997).
Figure 1.2: Structure of plant cell wall (Adapted from Pandya, 2011)
1.2.1 Cellulose:
Cellulose is the main constituent of plant cell wall comprising about 50% of wood and
closely associated with hemicelluloses and lignin. The basic structure of cellulose
consist of anhydroglucopyranoside units linked by β: 1-4 glycosidic bond. The
successive glucose residues are rotated by 180o relative to each other permit three
hydrogen bond per residue between each adjacent chain of cellulose and thus the
repeating unit of the cellulose chain is the cellobiose unit (Figure 1.3).
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Figure 1.3: Structure of cellulose (Adapted from Yang et al, 2007)
1.2.2 Hemicellulose:
The term hemicelluloses were first used to describe any plant polysaccharide that can
be extracted by mild alkali solutions (Yang et al, 2007). Hemicelluloses are generally
classified according to main sugar residue in the backbone, e.g. xylans, mannans,
galactans and glucans, with xylan and mannan being the main group of
hemicelluloses (Figure 1.4). Hemicellulose are often reported to be chemically
associated with or cross-linked to other polysaccharides, proteins or lignin. Xylans
appear to be the major interface between lignin and other carbohydrates.
Hemicelluloses are more soluble than cellulose, and they can be isolated from wood
by extraction. However, alkali extractions deacetylate the hemicelluloses completely.
The average degree of polymerization of hemicelluloses varies between 70 and 200
depending on the wood species (Fengel and Wegener, 1989).
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Figure 1.4: Structure of hemicellulose (Adapted from Yang et al, 2007)
1.2.3 Lignin
After cellulose lignin is the second most abundant renewable biopolymer in nature.
Lignin is an essential part of the plant cell wall, imparting rigidity and protecting the
easily degradable cellulose from attack by pathogens. Lignin is aromatic, three-
dimensional and amorphous. Lignin comprises of phenylpropanoid units joined
together by the action of peroxidase and laccase during lignin biosynthesis in the plant
cell wall (Boudet et al, 2003, Higuchi, 2006). Lignin is found in all vascular plants, a
major fraction being distributed throughout the secondary walls of woody cells and
also in the middle lamella between the secondary cell walls (Eriksson et al, 1990).
Lignin is a natural polymer with high molecular mass of up to 100 kDa or more
(Kästner, 2000) and can make up 20-30% of the lignocellulose in trees (Argyropoulos
and Menachem, 1997) there being a slightly higher content in gymnosperms
(softwoods) than angiosperms (hardwoods) (Eriksson et al, 1990). Lignin is deposited
as an encrusting and protecting material on the cellulose/hemicellulose matrix, and it
sets up a complex and acts as a kind of glue that cements the fibrous cell walls
together. Precursors of lignin synthesis are produced by plants from L-tyrosine and L-
phenylalanine which are synthesized from carbohydrates by the shikimic acid
metabolic pathway (Higuchi et al, 1977). They consist of an aromatic ring with up to
two methoxyl groups and three-carbon side chain designated as coumaryl, coniferyl
and sinapyl alcohol (Figure.1.5), yielding hydroxyphenol- (H-type), guaiacyl- (G-
type), and syringyl subunits (S-type) of lignin structure respectively (Higuchi, 1977).
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Figure 1.5: Basic structure of lignin (a) Structural unit of lignin; (b) Monomers of
lignin or monolignols (Adapted from Mendoza, 2011)
The ratio between syringyl and guaiacyl subgroups has been used as a comparative
parameter between plant species (Argyropoulos and Menachem, 1997). Guaiacyl
lignin is mainly found in softwoods (24 – 33% of dry biomass), guaiacyl-syringyl
lignin (16 – 25%) in hardwoods and grasses contain guaiacyl- syringyl- p
hydroxyphenol lignin (< 20%) (Sjöström and Wesrtermark, 1998). The methylation
of phenolic groups and thus the methoxyl content is recognized as an essential
criterion for lignin characterization (Brown, 1985). The O-methyl transferase is the
key enzyme in determining the composition of lignin. Gymnosperm, angiosperm, and
grass transferases catalyze different conversions leading to different precursors. This
explains the occurrence of different types of lignin and relates the O-methyl
transferases to the evolution of lignin.
The final step in lignin biosynthesis is peroxidase mediated dehydrogenation of the
phenyl propanoid precursors to produce phenoxyl radicals to yield large,
heterogeneous, and highly cross-linked polymer (Figure 1.6) (Eriksson et al, 1990).
The phenyl propanoid units are linked together through a variety of bonds, e.g. aryl-
ether,aryl-aryl, and carbon-carbon bonds (Adler, 1977). Lignin differs from other
natural polymers in that it has no single repeating bond (Brown, 1985). The
heterogeneity of this structure has been demonstrated through bindings of unusual
structures such as the dibenzodioxocin (Figure. 1.6) discovered by Brunow and
coworkers (Brunow, 2001). Due to this unique structure, lignin is highly resistant and
forms a barrier to microbial attack and degradation of wood.
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Figure 1.6: Structural model of lignin (Brunow, 2001)
1.3. Lignin modifying enzymes In nature, lignin is probably degraded by an array of microorganisms, although abiotic
degradation may also occur in environments, such as those due to alkaline chemical
spills (Blanchette, 1991) or UV radiation (Vähätalo et al., 1999). In aqueous or other
anaerobic environments, polymeric lignin is not degraded, and wood may persist in
non-degraded form for several hundred or thousand years (Blanchette, 1995).
Lignin degradation requires unspecific and extracellular enzymes because of the
random structure and high molecular mass (Kirt and Farrell, 1987). Most of the
studies conducted on laccases so far have focused on the model white rot species
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Trametes versicolor, Pleurotus ostreatus and a few others. However, the ecological
group of litter dwelling fungi (LDF), represented by species inhabiting the natural
environment of soil and decaying litter, are very promising candidates for the
ligninolytic activities. There are very few reports describing the presence of
ligninolytic activities in these species (Steffen et al, 2000 and 2003).
Lignin Degrading Enzymes (LDEs) belong to two classes, the heme containing
peroxidases and the copper containing laccases. The peroxidases comprises of
Manganese Peroxidase (MnP), Lignin Peroxidase (LiP) and Versatile Peroxidase
(VP). A series of redox reactions are initiated by the LDEs degrade lignin or the
structural analogous of the lignin subunits such as certain aromatic compounds. The
LDEs oxidize aromatic compounds until the aromatic ring structure is cleaved, which
is followed by further degradation with other enzymes. The principle characteristics
of these enzymes are depicted in Table 1.1.
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Table 1.1: Properties of Lignin modifying enzymes (Mendoza, 2011)
It requires H2O2 as its co-substrate and the presence of Mn +2 (Mn+2 is naturally
present in the wood) which oxidizes to Mn+3 and forms the Mn+3-chelate-oxalate
which in turn oxidizes the phenolic substrates.These chelates are small enough to
diffuse in to areas inaccessible even to the enzyme,as in the case of organopollutants
buried deep within the soil, which may not necessarily be available to the enzyme
(Hatakka ,2001).
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1.3.2 Lignin peroxidases (EC 1.11.1.14)
It too requires H2O2 as its co-substrate and the presence of mediator as a veratryl
alcohol to degrade lignin and other phenolic compounds. Here H2O2 gets reduced to
H2O by gaining an electron from LiP (which itself that gets oxidized). The oxidized
LiP then reduced to its native reduced state by gaining an electron from veratryl
alcohol and converting it to veratryl aldehyde. Veratryl aldehyde gets reduced to
veratryl alcohol by accepting an electron from lignin/pollutants. This results in the
oxidation of lignin or the analogous aromatic compounds (Hatakka, 2001).
1.3.3 Versatile Peroxidases (EC 1.11.1.16)
It is a novel enzyme which can utilize veratryl alcohol and Mn.+2 The most
noteworthy aspect of VP is that it combines the substrate specificity characteristics of
LiP, MnP and Cytochrome C Peroxidase. Hence it can oxidize wide variety of (high
and low redox potential) substrates including Mn +2, phenolic and non phenolic lignin
dimmers, veratryl alcohol, dimethoxy benzene, different types of dye, substituted
phenols and hydroquinones (Martinez, 2002 and Martinez et al, 2004).
1.3.4 Tyrosinase (EC 1.14.18.1)
It is an oxidoreductase enzyme, capable of acting as two types of catalysts using
molecular oxygen: monophenol oxidase and catecholase. As monophenol oxidase
(phenolase), it can oxidize phenols to o- quinones and produce o-diphenols, and as
catecholase (diphenolase), o-diphenols are oxidized to o-quinones. Some
biotechnological applications include biosensors for phenol, catechol and p-cresol
detection synthesis of antioxidants which is highly valuable in food applications,
removal of phenols in wastewaters and modification of polymers by addition of
quinones to chitosan (Mendoza, 2011).
1.3.5 Laccase (EC 1.10.3.2) –An important member of oxidoreductases
Laccases (benzenediol:oxygen oxidoreductases, EC 1.10.3.2), represents the largest
subgroup of blue multicopper oxidases (MCO), use the distinctive redox ability of
copper ions to catalyze the oxidation of a wide range of aromatic substrates
concomitantly with the reduction of molecular oxygen to water (Solomom et al, 1996
and Messerschmidt, 1997).
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1.4 Laccases in the natural environment Laccases are ubiquitous enzymes present in higher plants, bacteria, fungi, insects and
lichens (Riva, 2006; Lisov et al, 2007). The first report dates from 1883 when
Yoshida detected laccase-like activity in Rhus vernicifera (O´Malley et al, 1993).
However, laccase was designated as a p-diphenol oxidase in 1962 and accepted as
part of the lignification process in plants (O´Malley et al, 1993).
The considerable attention devoted to white-rot basidiomycetes and their ligninolytic
system in the past might lead to the conclusion that decaying wood is the most typical
environment for laccase production. Far less is known about the occurrence,
properties and roles of laccases occurring in other types of natural lignocellulose-
containing material like forest litter or soil. Plant litter is a major source of
lignocelluloses in the forest ecosystem. In the tropical rain forest the litter production
is 1.5 tons per hectare hence decomposition of plant litter by Litter
Dwelling/Decomposing Microorganisms (LDMs) is an important process of
controlling nutrient cycling and soil humus formation. Hence, if these communities
are not a part of the forest ecosystem we all would have been buried by the cast of
leaves and branches. Compared to wood, soil or litter is a more complex and
heterogeneous environment; hence soil or litter dwelling microorganisms are well
adapted to such competitive environment.
Relatively high activities of laccase – compared to agricultural or meadow soils – can
be detected in forest litter and soils in both broadleaved and coniferous forests, where
laccase is the dominant ligninolytic enzyme (Criquet et al, 2000 and Ghosh et al,
2003). Thus the presence of laccase activity reflects the course of the degradation of
organic substances. Laccase activity was found to increase during leaf litter
degradation in Mediterranean broadleaved litter (Fioretto et al, 2000) and the pattern
of detected isoenzymes varied during the succession (Nardo et al, 2004). In evergreen
oak litter, laccase activity was found to reflect the annual dynamics of fungal biomass
that is probably driven by the seasonal drying (Criquet et al, 2000).
Laccases as the most abundant ligninolytic enzymes in soil also attracted the
attention of ecologists studying its role in the carbon cycle, especially with respect to
the nitrogen input. Several studies documented a significant decrease of laccases and
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peroxidases in forest soils subjected to elevated nitrogen doses with the simultaneous
increase in the litter layer (Gallo et al, 2004). This phenomenon was accompanied by
the decrease of fungal biomass and the fungal: bacterial biomass ratio in soil as well
as by increased incorporation of vanillin as a model lignin-derived substrate into
fungal biomass; hence it seems that nitrate deposition directs the flow of carbon
through the heterotrophic soil food web (DeFerest et al, 2004). On the other hand, an
increase of phenolic compounds in forest soil after burning increased laccase activity
(Boerner & Brinkman, 2003). Similar to the situation in other lignocellulose-
containing substrates, laccases probably also participate in the transformation of
lignin contained in the forest litter. It is also generally presumed that laccases are able
to react with soil humic substances that can be directly formed from lignin
(Yavmetdinov et al, 2003). This is supported by the fact that humic acids induce
laccase activity and mRNA expression (Scheel et al, 2000). However, the interaction
of laccases with humic substances probably leads both to depolymerization of humic
substances and their synthesis from monomeric precursors; the balance of these two
processes can be influenced by the nature of the humic compounds (Zavarzina et al,
2004). Fakoussa & Frost, (1999) observed the decolorization and decrease of
molecular weight of humic acids, accompanied by the formation of fulvic acids
during the growth of T. versicolor cultures producing mainly laccase, and humic acid
synthesis was also documented in vitro using the same enzyme (Katase and Bollag,
1991).
Adsorption of laccases to soil humic substances or inorganic soil constituents changes
their temperature and activity profiles (Criquet et al, 2000) and inhibits its activity
(Claus and Filip, 1990).
1.4.1 Plants
The first laccase was reported in 1883 from as Rhus vernicifera, the Japanese lacquer
tree (Reinhammar, 1984), from which the designation laccase was derived. Laccases
have also been discovered from numerous other plants, for example sycamore (Bligny
and Douce, 1983), poplar (Ranocha et al, 1999), tobacco (De Marco and Roubelakis-
Angelakis, 1997) and peach (Lehman et al., 1974). In plants laccases are found in
xylem, where they presumably oxidize the monolignols in the early stages of
lignification (Gavnholt and Larson, 2002; Mayer and Staples, 2002; Bertrand et al,
Chapter 1 Introduction
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2002). Moreover, laccase have been shown to involve in the first step of healing of
wounded leaves (De Marco and Roubelakis- Angelakis, 1997).
1.4.2 Bacteria
The first reported laccase was found in Azospirrullum lipoferum, where laccase was
associated with the melanin production for cell pigmentation (Faure et al, 1994). In
other bacterial species, it was related with morphogenesis or the resistance of spores
against hydrogen peroxide and UV (Sharma et al, 2007). Characterization of bacterial
laccases has revealed that they have a low redox potential (0.45-0.54 V) but they are
active and stable at high temperatures (66 h at 60°C), pH (7-9) and salt
concentrations. These characteristics represent advantages for industrial applications,
since many processes are carried out under similar conditions where other types of
laccases might easily be inactivated (Durão et al, 2006).
1.4.3 Fungi – Source for an oxidative enzymes
Laccase activity has been demonstrated in several fungal species leading to the notion
that most of the fungi produce laccase. However, there are several physiological
group of fungi that apparently do not produce laccase. Laccase production has not
been demonstrated in lower fungi, that is in zygomycetes and chytridiomycetes
(Morozova et al, 2007). Several reports can be referred, in the literature on the
production of laccase in ascomycetes such as Gaeumannomyces graminis (Edens et
al, 1999) Magnoportha grisea (Iyer and Chattoo, 2003), Monocillium indicum
(Thakkar et al, 1992). In addition to plant pathogenic species, laccase production has
also been reported from some soil ascomycete species from the genera Aspergillus,
Culvularia, Penicillium, Fusarium (Banerjee and Vohra, 1991; Rodriguez et al, 1996.,
Scherer and Fischer, 1998; Chhaya and Gupte, 2010 and Mendoza, 2011) and in some
freshwater ascomycetes. The majority of laccases characterized so far have been
derived from white-rot basidiomycete fungi, which are efficient lignin degraders. A
well studied laccase producer white-rot fungi includes Coriolopsis rigida (Sapparat et
al, 2002); Fomes sclerodermeys (Papinutti et al, 2003); Phlebia radiata (Vares et
al,1995); Pleurotus ostreatus; Trametes hirsuta (Baldrian and Gabriel, 2002) and
Pleurotus pulmonarius ( De Souza et al, 2002).
Owing to the higher redox potential (+800 mV) of fungal laccases compared to plant
and bacterial laccases they are implicated in several biotechnological applications
Chapter 1 Introduction
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especially in the degradation of lignin and lignin related compounds (Bourbonnais et
al, 1995).
1.5 Properties and structure of laccases Current knowledge about the structure and physicochemical properties of fungal
proteins is based on the study of purified proteins. Up to now more than 100 laccases
have been purified from fungi and been more or less characterized. The laccase as a
holoenzyme form is a dimeric or tetrameric glycoprotein containing –per monomer
four copper atoms bound to three redox sites.Yoshida first discovered laccases in
1883 after an observation that latex from the Japanese lacquer tree (Rhus vernicifera)
hardened in the presence of air (Gianfreda et al, 1999). Laccases are defined in the
Enzyme Commission (EC) nomenclature as oxidoreductases which oxidize diphenols
and related substances and use molecular oxygen as an electron acceptor. Like most
enzymes, which are generally very substrate specific, laccases act on a surprisingly
broad range of substrates, including diphenols, polyphenols, different substituted
phenols, diamines, aromaticamines, benzenethiols, and even some inorganic
compounds (Xu, 1997). Laccases are multinuclear copper containing glycoproteins
that belong to the family of enzymes known as oxidases, more specifically “blue”
oxidases (Yaropolav et al., 1994) and phenol oxidases (Gianfreda et al, 1999).
Laccases from various sources vary greatly with respect to their degree of
glycosylation, molecular weight and kinetic properties (Yaropolav et al, 1994).
1.5.1 Molecular properties
Laccase is a glycosylated monomer or homodimer protein generally having fewer
saccharide compounds (10-25%) in fungi and bacteria than in the plant enzymes. The
carbohydrate compound contain monosaccharide such as hexosamines, glucose,
mannose, galactose, fructose and arabinose (Rogalski and Leonowicz, 1991). On
SDS-PAGE, most laccases show mobilities corresponding to molecular weight of 60-
100 kDa, of which 10-50% may be attributed to glycosylation. Mannose is one of the
major components of the carbohydrate susceptibility, activity, copper retention, and
thermal stability (Xu.,1999). However two domain laccases with molecular weight of
30 and 40 kDa have been isolated from Botrytis cinerea (Nakamura and Go, 2005)
and fresh fruiting bodies of Trichoderma giganteum (Wang and TB, 2004).
Chapter 1 Introduction
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Most laccases studied are extracellular protein, although intracellular laccases have
been detected in several fungi and insects. Fungal laccases have isoelectric point (pI)
ranging from 3.0 to 7.0, whereas plant laccases pI value range to 9.0. The main
difference between the two enzymes is that fungal enzymes have their pH optima
between 3.6 to 5.2, while laccase from Rhus venifera have pH optima between 6.8 to
7.4. The low pH optima may be because of their adaptation to grow under acidic
conditions, while the plant laccases being intracellular having pH optima nearer to
their physiological pH (Madhavi and Lele, 2009).
1.5.2 Spectral properties
Laccases generally exhibit two absorption peaks when subjected to U.V.Visible
wavelength scan, a strong absorbance is visible at 600 nm and is associated with the
type-1 copper, while the shoulder at 330 nm is an indicative of the type-3 pair of
copper atoms. There are reports such laccases that do not display this characteristic
spectrum. A “white” laccase was said to be isolated from Pleurotus ostreatus
(Palmieri et al, 1997), while Leontievsky et al, (1997) reported the presence of
“yellow” laccases. The loss of the absorption peak at 600 nm of the “white” laccase
was attributed to the presence of only a single copper atom in the metal cluster, the
other three atoms being replaced by two zinc and one iron atom (Palmieri et al, 1997).
Leontievsky et al, (1997) showed that the loss of this peak in the case of “yellow”
laccases to copper atoms being present in their reduced state.
For the catalytic mechanism minimum of four copper atoms per active protein unit is
needed (Lentievsky et al, 1997). The type 1 copper (T1) is responsible for the intense
blue colour of the enzyme and has a strong electronic absorption around 600nm and is
EPR detectable. The type 2 copper (T2) is colourless but EPR detectable, and type 3
copper (T3) consists of a pair of copper atoms that give weak absorbance near the UV
spectrum but no EPR signal. The T2 and T3 copper sites are close together to form
and form a trinuclear centre (Lentievsky et al, 1997) in which binding of dioxygen
and four electron reduction to water occur. Not all laccases are reported to possess
four copper atoms (Thurston et al, 1994) per monomeric molecule.
Chapter 1 Introduction
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1.5.3 Substrate specificity
Laccase (EC 1.10.3.2) is a blue copper protein, but also falls within the broader
description of polyphenol oxidases. Polyphenol oxidases are copper proteins with the
common feature that they are able to oxidize aromatic compounds with molecular
oxygen as the terminal electron acceptor (Mayer, 1987). Polyphenol oxidases are
associated with three types of activities:
Catechol oxidase or o-dipenol: oxygen oxidoreductase (EC 1.10.3.1)
Laccase or p-diphenol: oxygen oxidoreductase (EC 1.10.3.2)
Cresolase or monophenol monooxygenase (EC 1.18.14.1)
There is, however, difficulty in defining laccase according to its substrate specificity,
because laccase has an overlapping range of substrates with tyrosinase. Catechol
oxidases or tyrosinases have o-diphenol as well as cresolase activity (oxidation of L-
tyrosine). Laccases have ortho and paradiphenol activity, usually with more affinity
towards the second group. Only tyrosinases possess cresolase activity and only
laccases have the ability to oxidize syringaldazine (Thurston, 1994; Eggert et al,
1996). Laccases are remarkably nonspecific as to the inducing substrate, and the range
of substrate oxidized varies from one laccase to another (Wood, 1980).
1.5.4 Isozymes of laccase
A single organism may also possess several laccase isozymes (or isoforms) that may
differ in their amino acid sequence and display different kinetic properties towards
standard laccase substrates.Many laccase producing fungi secrete isoforms of the
same enzyme. These isozymes have been found to originate from the same or
different genes encoding for the laccase enzyme. The number and of isoforms vary
with species and also within species. The biochemical characteristics of isoezymes
vary depending upon the source and culture conditions (Desai and Nityanand, 2011).
Two laccase isoenzymes (POXA1 and POXA 2) produced by Pleurotus ostreatus
with molecular weight of 61 and 67 kDa, pI of 6.7 and 4 respectively (Palmieri et
al,1997). Four laccase isoenzymes (LCC1, LCC2, LCC3 and LCC4) synthesized by
Pleurotus ostreatus strain V-184were purified and characterized (Mansur et al, 2003).
LCC1 and LCC2 have molecular masses of about 60 and 65 kDa and exhibited same
pI value of 3.0. Laccases LCC3 and LCC4 were characterized by SDS PAGE,
estimating their molecular masses around 80 and 82 kDa, pI 4.7 and 4.5 respectively.
Chapter 1 Introduction
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When staining with the ABTS and guaiacol in native polyacrylamide gels, different
specificities were observed for LCC1/LCC2 and LCC3/LCC4 isoenzymes. Three
laccase isoenzymes Lac1, Lac2 and Lac3 from C.unicolor had significantly varying
biochemical characteristics (D’Souza-Ticlo et al, 2009).
1.5.5 Structural properties
Three-dimensional structural analysis of several fungal, bacterial and plant laccases
reveals that all are composed of three sequentially arranged domains; each of them
with a greek key β-barrel topology, highly related to small copper proteins such as
azurin and plastocyanin.(Giardina et al, 2010). The multiple alignment of primary
sequences of laccases shows that the copper binding motifs are highly conserved in all
sequences, which reflects a common mechanism for copper oxidation and oxygen
reduction. However, putative binding pocket analysis reveals that bacterial laccases
have larger binding cavities when compared to those from plants and fungi (Mendoza,
2011). Generally, laccase contains four copper atoms (Figure 1.7), which have been
classified into three groups based on the absorption and Electronic Paramagnetic
Resonance spectra. Type 1 (T1) paramagnetic “blue” copper has an intense absorption
at 600-610 nm, which is caused by the covalent copper-cysteine bond and confers the
typical blue color to the multicopper proteins. The T1 copper has a trigonal
coordination with two histidines and one cysteine; in bacterial laccases the axial
ligand is conformed by methionine and in fungal laccase by leucine or phenylalanine
(Witayakran & Ragauskas, 2009). Type 2 (T2) paramagnetic “non-blue” copper has
no visible absorption spectrum and is coordinated by two histidines. Type 3 (T3) is a
diamagnetic coupled binuclear copper center, with an absorption band at 330 nm. It is
coordinated by six histidines (Claus 2004; Witayakran & Ragauskas,2009).
Nevertheless, it is possible to find non-blue laccases in nature (Palmieri et al 1999);
the “white” laccases, as they are called, have been structurally characterized and
atypically show the presence of one copper, one iron and two zinc atoms per
molecule.
Chapter 1 Introduction
18
Figure 1.7: Schematic representation of Copper centers from fungal laccase (Claus, 2004)
Structural analysis of Trametes versicolor laccase and site-directed mutagenesis in
Bacillus sp. laccase have revealed that the axial ligand in T1 copper is responsible for
displaying the redox potential; T1 copper has no axial ligand in Trametes versicolor
laccase and this has given rise to a modest elevation of its redox potential to 0.78 V
(Piontek et al, 2002). Moreover, mutations of Bacillus sp. laccase have been used to
confirm that modifications in the axial ligand of T1 (methionine was replaced by
phenylalanine or leucine) allowed changes in the redox potential (the change of amino
acids led to an increase of 0.06-0.1 V of the redox potential as compared to the wild
type) (Durão et al, 2006). The redox potential is directly related to how good a laccase
will catalyze oxido-reduction reactions.
Different compounds have been reported as inhibitors of laccase. Among them,
anions like azide, cyanide and fluoride inhibit laccase by binding T2/T3, thus
preventing electron transfer from T1. Other inhibitors like metal ions, fatty acids and
quaternary ammonium detergents replace or chelate the copper centers and may also
denature the protein (Witayakran & Ragauskas, 2009).
Chapter 1 Introduction
19
1.6 Catalytic mechanism Laccase only attacks the phenolic subunits of lignin, leading to Cα oxidation, Cα-Cβ
cleavage and aryl-alkyl cleavage. The substrate oxidation by laccase is a one electron
reaction generating a free radical. The product formed initially is unstable and may
undergo second enzyme catalyzed oxidation or a non-enzymatic reaction such as
hydration, disprotonation or polymerization. Thus laccase can be thought to operate as
a battery, storing electrons from individuals oxidation reactions in order to reduce
molecular oxygen. Hence the oxidation of four reducing substrate molecules are
necessary for the complete reduction of molecular oxygen to water (Thurston, 1994).
Laccase are known to reduce wide range of aromatic compounds which includes
polyphenols methoxy-substituted monophenols and aromatic amines (Bourbonais et
al, 1995).
The catalytic cycle of laccase and its proposed mechanism for the reduction and
reoxidation of Cu+2 sites is illustrated in figure 1.8. In this the substrate reduces the
water reduces the T1 site, in the “native intermediate” which than transfers the
electron to the trinuclear cluster T2/T3. The T1 and T2 sites together reduce T3 pair,
and each copper in the cluster is sequentially reduced by electron transfer from the
T1site, in this case the T3 site no longer acts as a two electron acceptor. Ultimately
the slow decay of the “native intermediate” leads to the formation of the resting, fully
oxidized form, which will culminate the catalytic cycle with the reuction of oxygen to
water (Witayakran & Ragauskas, 2009).
Chapter 1 Introduction
20
Figure1.8:Mechanism of four-electron reduction of molecular oxygen to water in the catalytic cycle of laccase. (Modified from Witayakran & Ragauskas 2009)
The catalytic mechanism of laccase can be summarized in three steps:
(i) Type-1 copper reduction by the reducing substrate,
(ii) Internal electron transfer from type 1 copper to type 2 and type 3 copper
trinuclear cluster,
(iii) Molecular oxygen reduction to water at type-2 and type-3 copper atoms.
The structure of laccase active site showing the flow of substrate, electron and oxygen
is shown in figure 1.9.
Chapter 1 Introduction
21
Figure 1.9: The structure of the laccase active site with arrows marking the
flow of substrates, electrons (e-) and O2. (Solomon et al, 2008)
1.6.1 Natural substrates
The natural substrates of laccase include phenols like ortho- and paradiphenols,
aminophenols, polyphenols, polyamines and aryl diamines. The oxidation of these
molecules is represented in figure 1.10. Here, laccase oxidizes the molecule with a
simultaneous radical formation, which can spontaneously rearrange to cleave the
aromatic rings or promote their polymerization.These phenolic compounds are typical
substrates for laccase due to their low redox potential (0.5-1 V); however, other non-
phenolic structures (including some phenolic compounds) might have a higher redox
potential, which determines the low efficiency of laccase towards the substrate (Couto
& Toca-Herrera, 2006).
Chapter 1 Introduction
22
Figure 1.10: Oxidation of phenolic compounds (natural substrates) by laccase (Madhavi and Lele, 2009)
1.6.2 Laccase/Mediator system
Laccase has the ability to oxidize only phenolic fragments of lignin due to the random
polymer nature of lignin and to its low redox potential. However, small natural low
molecular weight compounds with high redox potential than laccase called
“mediators” may be used to oxidize the nonphenolic part of lignin. These “mediators”
extend the oxidation potential of laccase and have the capacity to change the redox
potential during the oxidation(Bourbonnais et al, 1998; Zille et al, 2003). i.e. from
0.78 V to 1.084 V (Zille et al, 2003). Consequently, the oxidation of non-natural
substrates and/or with high redox potentials (like lignin) is possible. The catalytic
mechanism involves the oxidation of the mediator, which can diffuse away from the
enzyme, oxidize the substrate and return to the catalytic cycle as a reduced species
(Riva, 2006). Mediators can oxidize the substrate by different mechanisms; those
containing N-OH oxidize the substrate via hydrogen atom transfer pathway, whereas
others (i.e., ABTS) do it via electron transfer [Figure1.11 (a), (b), (c)].
Chapter 1 Introduction
23
Several organic and inorganic compounds have been reported as effective mediators
In 1990, the diammonium salt 2,2'-azinobis (3-ethylbenzothiazoline-6-sulfonic acid)
(ABTS) was found to mediate and enhance the laccase activity. Later,
Nhydroxyphthalimide (HPT) and 2,2,6,6-tetramethylpiperidine-N-oxyl (TEMPO)
(and its derivatives) were reported to act as mediators (Morozova et al, 2007). The
latter was proven to be the most effective towards lignin degradation The application
of these mediators can be limited due to their high cost as well as their toxicity.
Nevertheless, this can be overcome by the application of immobilized mediators,
which allows their recyclability and facilitates their disposal by using membranes that
retain the immobilized molecule (Kunamneni et al 2007). On the other hand,
molecules like vanillin, p-coumaric acid, acetovanillone, methyl vanillate,
syringaldehyde, and some dyes are reported as “natural” mediators. They have been
shown to catalyze reactions as effectively as the other type of mediators (Camarero et
al, 2007). Additionally, they can be easily produced from lignin (Camarero et al,
2005).
The use of molecular oxygen as the oxidant and the fact that water is the only by-
product are very attractive catalytic features, rendering laccases as excellent ‘green’
catalysts.
Chapter 1 Introduction
24
Figure 1.11: Laccase-mediated oxidation of substrates in the presence of synthetic and natural mediators. ArCH2OH is the substrate and represents an aromatic alcohol. a) A type of N-OH as the synthetic mediator showing the H-atom transfer pathway; b) a synthetic mediator showing the electron transfer pathway; c) a phenolic compound as the natural mediator. (Adopted from Mendoza , 2011)
The role of laccase in PAH degradation has been well studied. Laccase can also
catalyze one electron oxidation of PAHs such as anthracene and benzo[a]pyrene that
both have ionization potentials 57.55 eV. Although the PAH oxidizing ability of
white-rot fungi Pleurotus ostreatus closely correlates with its laccase activity and
lignin degrading ability. However, it was later shown that laccase has a role in PAH
oxidation by WRF. Crude enzyme preparations as well as two purified isoenzymes
from Trametes versicolor were able to oxidize anthracene and benzo[a]pyrene
(Collins et al, 1996). Direct oxidation of anthracene by the two purified laccases with
ABTS was observed but a marked increase in levels of oxidation occurred when
present (Collins et al, 1996). In contrast, no significant direct oxidation of
Chapter 1 Introduction
40
benzo[a]pyrene by purified laccase was observed. The presence of ABTS in the
reaction mixture was essential for high levels of benzo[a]pyrene oxidation (Collins et
al, 1996). The activity of laccase is restricted to compounds with low ionization
potentials such as aromatic compounds with a phenolic functional group (Bohmer et
al., 1998). The substrate range of laccase extends to nonphenolic lignin structures
when mediating substrate compounds such as ABTS are present. The mediating
substrate presumably functions as a diffusible redox mediating substrate between that
compound and the enzyme. It has been demonstrated that such laccase/mediating
substrate couples oxidize PAHs and that the IP threshold value for the oxidation of
PAHs by laccase appears to be similar to that of LiP (Bohmer et al, 1998).
1.10.2.8.2 Alkanes
Laccase from the white-rot fungus, Trametes hirsuta,has been used to oxidize alkenes.
The oxidation is the effect of a two-step process in which the enzyme first catalyzed
the oxidation of primary substrate, a mediator added to the reaction, and then the
oxidized mediator oxidizes the secondary substrate, the alkene, to the corresponding
ketone or aldehyde. The best results were obtained by using hydroxybenzotriazole as
mediator, and aliphatic polyunsaturatedand aromatic allyl alcohols were completely
oxidized within 2 h at 20 ºC. Aliphatic allyl alcohols were oxidized up to 70% at 45
ºC for 20 h. By contrast, the oxidation of other alkenes, such as allyl ether, cis-2-
heptene and cyclohexene, did not exceed 25% (Niku and Viikari, 2000).
1.10.2.8.3 Dyes
Laccase purified from different fungi, were able to degrade triarylmethane, indigoid,
azo, and athraquinonic dyes used in dyeing textiles. Immobilization of the T. hirsuta
on alumina enhanced the thermal stabilities of the enzyme and its tolerance against
inhibitors such as halides, copper chelators, and dyeing additives. Treatment of the
dyes with immobilized laccase reduced their toxicity up to 80% based on oxygen
consumption rate of Pseudomonas putida. Textile effluents decolorized with
immobilized laccase could be used for dyeing, and acceptable colour differences were
measured for most dyes (Abadulla et al, 2000).
Chapter 1 Introduction
41
1.10.2.8.4 Industrial wastes
An isolate of the fungus, Flavodon flavus, was shown to be able to decolourize the
effluent from a Kraft paper mill bleach plant. The important enzymes, produced by
the fungus in the presence of the effluent, were laccase, manganese-dependent
peroxidase, and lignin peroxidase. The culture appeared to be a potential candidate for
bioremediation of coloured industrial effluents (Raghukumar, 2000).
1.10.2.8.5 Herbicide
Isoxaflutole is an herbicide activated in soils and plants to its diketonitrile derivative,
the active form of the herbicide. The diketonitrile derivative undergoes cleavage to
the inactive benzoic acid analogue. Laccase enzymes in two fungi, Phanerochaete
chrysosporium and T. versicolor, are able to convert the diketonitrile to the acid, as
will purified laccase in the presence of 2 mM 2,2-azinobis(3-ethylbenzthiazoline- 6-
sulfonic acid) acting as a redox mediator at pH 3 (Mougin et al, 2000).
1.10.2.8.6 Phenolic environmental pollutants –Bisphenol A
Bisphenol A [BPA: 2, 2-bis (4-hydroxyphenyl)propane] is widely used in a variety of
industrial and residential applications such as the synthesis of polymers including
polycarbonates, epoxy resins, phenol resins, polyesters, and polyacrylates. The
chemical structure of BPA consists of two phenolic rings joined together through a
bridging carbon. Recently, biphenolic compounds including BPA have been
recognized as endocrine disrupting chemicals (EDCs). (Sajiki et al, 2002).
Bisphenol A, commonly abbreviated as BPA, is an organic compound with two
phenol functional groups. It is used to make polycarbonate plastic and epoxy resins,
along with other applications (Figure 1.13).
Figure 1.13: Synthesis of Bisphenol A by the condensation of acetone and two equivalents of phenol. (Adapted from wikepedia).
BPA is suspected to reduce the number of sperms in men and to act as a risk factor for
the development of prostate and breast cancer (Kang et al, 2006a; 2006b). Biological
Chapter 1 Introduction
42
effects in aquatic animals have been reported to develop from 1- 10 mg/l of BPA
upwards, including effects on the sex ratio and fertilization (Maffini et al, 2006). BPA
is a plasticizer of polycarbonate plastics, which are used for food packaging, coatings
of metal cans and baby bottles. BPA can leach out from these materials during
washing and sterilization processes or after landfilling (Coors et al, 2003; Krishnan et
al, 1993). Thus, it is necessary to assess its biodegradability or fate in the natural
environment. Several research groups reported the biodegradation of BPA using
enzymes from lignin-degrading basidiomycetes.
A number of microorganisms is capable of degrading or metabolizing BPA, among
them white-rot fungi (Kang et al, 2006a) actually colonizing wood or leaf-litter. They
produce and secrete highly active oxidative enzymes, of which extracellular laccase
and manganese peroxidase (MnP) are the most common ones. These enzymes have
been shown to degrade a variety of man-made organopollutants (Steffen et al, 2003;
Scheibner & Hofrichter, 1998). Among white-rot fungi, the ecophysiological group of
litter-decomposing fungi appears to have additional interesting properties for
bioremediation purposes. They are able to co-exist and also compete with the
indigenous soil microflora, since grasslands and topsoil layers are the natural habitats
of these fungi. Thus, litter-decomposing fungi were selected to study the conversion
of BPA and the removal of its estrogenic activity.
These results strongly suggested that ligninolytic enzymes were effective for the
removal of the estrogenic activity via the oxidative degradation of BPA. Although the
degradation of BPA by ligninolytic enzymes was reported, this process has been
attempted in aqueous media with a limited concentration of the pollutant. In general,
environmental pollutants such as BPA and p-nonylphenol do not dissolve in aqueous
media owing to their high hydrophobicity and hence organic solvents are required to
dissolve them. This implies that the use of organic solvents inevitably allows the
degradation reaction to proceed at a high concentration of environmental pollutants
and in a homogenous system. However, native enzymes do not exhibit significant
catalytic activities in organic media.
Chapter 1 Introduction
43
1.11 Potential new laccase based biocatalysis-Non aqueous approach
Classical “in water enzymology” usually deals with “purified enzyme preparations
that can lack on isolation from living matter some of the components essential for
exposition of real catalytic properties in vivo. In nature enzyme function in
microheterogenous systems, for example interacting with different surfaces composed
from lipid membranes or being incorporated in to biomembranes. Even in cytoplasm,
water is not a dominating component and is playing a structural role as well by
participating in formation of biocatalytic complexes (generally of glycolipoprotein
origin) (Levashow, 1992). During the past decade much progress has been made in
fundamental understanding of the phenomena that govern biocatalysis in non-
conventional media. The factors that affect biocatalytic reactions and the activity and
stability of biocatalysis in these reaction media are generally associated with the
crucial role of water and the need to keep biocatalysis in active conformation
(Vermuë and Tramper, 1995).
The non-conventional media deal with the use of organic solvents and supercritical
fluids. There are several potential advantages for the introduction of organic solvents
in synthetic reactions.
1. Organic solvents will increase the solubility of poorly water-soluble substrates,
thereby improving the volumetric productivity of the reaction.
2. The thermodynamic reaction equilibrium may be shifted to favor synthesis over
hydrolysis, either by altering the partitioning of the substrate/product between the
phases of interest, or by reducing the water activity.
3. Higher product yields can be achieved by reduction of substrate and/or product
inhibition, either indirectly by maintaining a low concentration in aqueous micro-
environment of the biocatalyst (Schwartz and McCoy, 1977; Vermuë and Tramper,
1990), or directly by changing the interaction between the inhibitor and active site of
the enzyme (Zaks and Russel, 1988).
4. Application of low boiling point solvents will simplify recovery of the product and
the biocatalyst.
5. Thermostability of the enzymes is improved when microaqueous reaction media are
used (Zaks and Kibnov, 1984; Volkin et al, 1991).
Chapter 1 Introduction
44
6. Possibility to manipulate the stereo-and regio-selectivity of the enzyme in such media
(Sakurai et al, 1988).
Four categories of organic solvent reaction media for biocatalysis can be
distinguished. The water/organic solvent mixtures may consist mainly of water with
relatively small amount of water miscible solvents (Figure 1.14A). The mixture may
consist of a two phase system of a water –immiscible organic solvent and an aqueous
buffer (Figure 1.14 B1,B2,B3) or it may be an organic solvent in which dry
biocatalyst is suspended, so- called microaqueous organic-solvent mixture (Figure
1.14C). The fourth category of organic-solvent reaction media is the reverse micells
(Figure 1.14 D). Reverse micells consist of tiny droplets of aqueous medium (radii in
the range of 1-50 nm) stabilized by surfactant in a bulk of water-immiscible organic
solvent.
Figure 1.14: Scematic representation of the four categories of organic-solvent reaction media. A: Water misscible solvent. B1: Two phase system, low volume organic solvent, solubalized biocatalyst. B2: Two phase system, low volume organic solvent, immobilized bio catalyst. B3: Two phase system, aw= 1, high volume organic solvent, immobilized biocatalyst. C: Micro-aqueous system, aw < 1 D: Reverse micelles. (Adopted from Vermuë and Tramper, 1995)
Chapter 1 Introduction
45
1.11 Aim and scope of present investigation Laccase produced by different microorganisms was found to be promosing candidate
for bioremediation of variety of recalcitrant compounds including phenolic pollutants.
Endocrine disrupting chemicals (EDCs) are naturally occurring compounds or man-
made chemicals that act like hormones in the endocrine system and disrupt the
physiologic function of endogenous hormones. Bisphenol A (BPA), known as one of
EDCs since 1936 and has aroused the public concerns. The potential adverse effects
of BPA on human health and reproductive biology include breast and prostate cancer;
sperm count reduction, abnormal penile/urethra development in males, early sexual
maturation in females, neurobehavioral problems, prevalence of obesity, type 2
diabetes and immunodeficiency. A number of methods such as electrochemical
process, sonochemical degradation, ozonation, chemical oxidation photooxidation
solvent extraction membrane filtration, and sorption have been employed to eliminate
Bisphenol A from wastewater. These processes are costly, less efficient and not
environmental friendly. Laccase based bioremediation of phenolic pollutant like
Bisphenol A can serve as a more environmentally benign alternative. The present
research work is aimed to search for an efficient laccase producing sysem followed by
its optimization, strain improvement, scale-up and application in organic media for
the bioremediation of phenolic environmental pollutant Bisphenol A.
The following are the broad objectives of the present investigation
Isolation of novel laccase producing microorganism, its characterization and
optimization of physiological conditions for laccase production.
Statistical optimization of medium constituents for higher production of laccase.
Strain improvement to increase laccase productivity.
Scale-up and evaluation of improved strain for laccase production under solid
substrate tray fermentation.
Purification and characterization of laccase from the laccase hyper producing strain.
To study the efficacy of purified laccase in reverse micelles system for the
degradation of phenolic environmental pollutant Bisphenol A.
Chapter 1 Introduction
46
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